Articles in PresS. Am J Physiol Heart Circ Physiol (January 16, 2003). 10.1152/ajpheart.01008.2002
FIBROBLAST ALIGNMENT UNDER INTERSTITIAL FLUID FLOW
USING A NOVEL 3-D TISSUE CULTURE MODEL
Chee Ping Ng1 and Melody A. Swartz1,2
Departments of Chemical Engineering1 and Biomedical Engineering2,
Northwestern University, Evanston, Illinois 60208
Address for correspondence:
Melody A. Swartz, Ph.D.
Department of Biomedical Engineering
Northwestern University
2145 Sheridan Rd
Evanston, IL 60208-3107
Tel: (847) 467-6668
Fax: (847) 491E-mail: [email protected]
Running title: Interstitial flow in vitro
Copyright (c) 2003 by the American Physiological Society.
Abstract
Interstitial flow is an important component of the microcirculation and interstitial
environment, yet its effects on cell organization and tissue architecture are poorly
understood, in part due to the lack of in vitro models. To examine the effects of
interstitial flow on cell morphology and matrix remodeling, we have developed a tissue
culture model that physically supports soft tissue cultures and allows microscopic
visualization of cells within the 3-D matrix. In addition, pressure-flow relationships can
be continuously monitored to evaluate the bulk hydraulic resistance as an indicator of
changes in overall matrix integrity. We observed that cells such as human dermal
fibroblasts aligned perpendicular to the direction of interstitial flow. In contrast,
fibroblasts in static 3-D controls remained randomly oriented while fibroblasts subjected
to fluid shear as a 2-D monolayer regressed. Also, the dynamic measurements of
hydraulic conductivity suggest reorganization towards a steady state. These primary
findings help establish the importance of interstitial flow on the biology of tissue
organization and interstitial fluid balance.
Key words: Cell culture, mechanical stress, shear stress, hydraulic conductivity
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Introduction
Physiological interstitial flow is the movement of fluid through the extracellular
matrix of a tissue, often between blood vessels and lymphatic capillaries. It provides
convection necessary for the transport of large proteins through the interstitial space and
constitutes an important component of the microcirculation (Fig. 1). Aside from its role
in transport, interstitial flow also provides a specific mechanical environment to cells in
the interstitium that could play an important role in determining interstitial organization
and architecture, based on abundant evidence that mechanical forces help govern the
architecture of tissues such as lung (16, 33), bone (1, 11, 13), articular cartilage (8, 20),
and vascular tissues (3, 4, 17, 23, 25). Importantly, many cell types including fibroblasts,
smooth muscle cells, osteoblasts and chondrocytes reside within a 3-D environment and
are exposed to interstitial fluid forces; this is in contrast to cells such as endothelial and
epithelial cells that form a monolayer to create a lumen or surface and are exposed to
shear stresses across the surface.
Despite its importance, the biological regulation of interstitial fluid balance is
poorly understood, largely because of the lack of experimental models. Although many
in vivo and in vitro studies have been performed to characterize the mechanics of
interstitial fluid balance and estimate the Darcy permeability of a variety of tissues, there
have been very few studies to examine how interstitial flow affects cell response and how
interstitial fluid is regulated in a soft tissue environment. In their seminal study, Wang
and Tarbell investigated the effects of transvascular flow on smooth muscle cells seeded
in a collagen gel model and found that the production levels of prostaglandins were
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tenfold lower in cells under interstitial fluid shear than in a 2-D monolayer shear model
using a rotating disk (30), demonstrating that cell response to fluid flow in 2-D
configurations poorly mimics the response of fluid flow on interstitial cells in their
natural 3-D environment. Furthermore, it has been demonstrated that medium perfusion
improves the architecture of engineered tissues such as cardiac muscles (2),
downregulates extracellular protein accumulation in engineered cartilage (19), and
enhances the viability and functionality of bone cells seeded in collagen constructs (7,
21).
In this study we develop an in vitro flow chamber to directly examine in situ the
effects of interstitial flow on cell organization in soft tissue cultures over relatively long
periods of time (days to weeks), where we can (i) specifically control the flow
environment, (ii) directly observe cell organization, and (iii) measure the mechanical
properties (e.g. hydraulic resistance) ‘online’. In doing so, we observe that fibroblasts
align perpendicular to interstitial flow in 3-D culture. These novel findings, along with
the model we developed, help to form an experimental basis for understanding the
biological regulation of interstitial fluid balance in soft tissues.
Methods
Cell culture. Human dermal fibroblasts (CCD1079sk, ATCC, Manassas, VA)
were cultured in alpha-modified Minimum Essential Medium Eagle (α-MEM; SigmaAldrich, St. Louis, MO) with 10% fetal bovine serum (GIBCO, Carlsbad, CA).
Flow model set-up and preparation. The interstitial flow chamber consisted of a
cell-suspended collagen gel sandwiched between two glass coverslips, 1.6 mm apart,
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through which a radial flow of culture medium was directed (Fig. 2). The gel was affixed
between the flow inlet and outlet by acid-treated porous polytheylene (PE), which
anchored the collagen and prevented contraction (14).
To enhance binding of the
collagen gel to the top and bottom glass surfaces, they were cleaned (26), silanized with
2% 3-Aminopropyltriethoxysilane (Pierce, Rockford, IL) for 15 minutes, and
functionalized with 0.1% glutaraldehyde for 30 minutes before rinsing thoroughly with
deionized water. The components of the flow chamber were assembled under sterile
conditions with sterile silicone glue.
After chamber assembly, the chamber was filled with a cell-populated 0.35%
collagen solution. The collagen solution was prepared from rat tail tendons according to
Pasternak and Miller (22). Human dermal fibroblasts were added to the collagen at a
density of 105 cells/ml. The cell-gel solution (of approximately 150 µl) was injected into
the chamber and incubated at 37°C for 30 minutes to allow the gel to polymerize and
react with activated glass surfaces. The entire setup was then placed in a 35mm plate and
immersed in media overnight for cell attachment in a 37°C / 5% CO2 incubator.
To induce flow, the chamber was connected to a sterile media reservoir via a
peristaltic pump and a pressure manometer.
The flow was delivered at 10 µl/min
(leading to an average velocity of 13 µm/s at the inlet and 3.6 µm/s at the outlet, and an
average residence time of approximately 14.5 min), which induced an average pressure
drop of 3.5 cm H2O. A static chamber used as a control was set up exactly as the
experimental flow chamber, except that it was not connected to the flow delivery
apparatus. In both groups, medium surrounded the chamber and could diffuse through
the outer and inner PE rings; this medium was replaced every 2 days. All cultures were
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maintained in a humidified 37°C, 5% CO2 incubator for the duration of the experiment.
The manometer readings were monitored every 3 hours and the culture system was
examined under light microscopy (Nikon TE 200) daily.
2-D monolayer shear studies. To compare the effects of interstitial flow with
flow across a 2-D monolayer, we used the same flow chamber set-up as the interstitial
flow studies, but instead of filling the chamber with cell-suspended gel, we seeded the
cells at a density of 104 cells/cm2 atop a thin coating of 0.35% collagen that was
covalently bound to the bottom functionalized surface of the chamber. After 24-48 hours
of static culture to allow the cells to attach to the surface, at which time the cells reached
a confluence of 30-40%, flow was induced over the cell monolayer for 2 days under the
same volumetric flow rate as in the 3-D cultures (10 µl/min).
Immunofluorescent staining. Fluorescein phalloidin (Molecular Probes, Eugene,
OR) was used to examine the alignment of the cells when subjected to interstitial flow by
staining its actin fibers.
The entire chamber was fixed by immersion in 4%
paraformaldehyde in PBS for 30 minutes, rinsed with PBS, and permeabilized in 0.5%
Triton for 30 minutes.
The chamber was then immersed overnight in a solution
containing 200nM fluorescein phalloidin. The gels were rinsed in PBS and images were
taken with epi-fluorescence microscopy (Nikon TE 200 with SPOT RT Slider digital
camera) and laser scanning confocal microscopy (Leica LCS Laser Microscope System).
For the flow studies on fibroblast monolayers (used for comparison of cell behavior in 3D gels), the fibroblasts were live-labeled with 5 µM Calcein AM (Molecular Probes) for
10 minutes prior to the induction of flow. This calcein AM live stain was unsuitable for
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long-term use in the 3-D studies since the dye was not retained in viable cells after
several cell divisions (10).
Computation of bulk Darcy permeability. Continuous online measurements of
overall Darcy permeability K′, or the inverse hydraulic resistance, were used as a bulk
indicator of changes in matrix integrity. K′ (cm2) was evaluated according to Darcy’s law
in a 1-D radial configuration where K′=[Qµ ln(ro/ri)]/[2πh∆P(t)] and Q is the constant
volumetric flow rate delivered by the peristaltic pump, µ is the viscosity of the perfusing
fluid (=1.2 cp), h is the gel thickness (= 1.6 mm), ∆P(t) is the pressure gradient driving
the flow (i.e. the difference between the inlet pressure as measured from the manometer
and the outlet pressure of 1.082 cm H2O, equal to the height of medium in the dish), and
ro and ri are the outer and inner radii of the gel, respectively (=5.6 mm and 1.6 mm).
(Note: the hydraulic conductivity K (cm4/dyn/s) is also commonly used in the literature,
where K=K′/µ.) The collagen gel dominated the overall hydraulic resistance since the PE
had a value of K′ that was two orders of magnitude greater than that of the gel (10-4 vs.
approx. 10-6 cm2 respectively). Also, the Reynold’s number was estimated at 10-5 to 10-6
for this system, validating the use of Darcy’s law in modeling interstitial flow through the
gel culture system.
Measurement of cell angle.
Cell orientation in the composite radial section
micrographs was quantified for representative flow and static conditions using Adobe
Photoshop 6.0 (San Jose, CA) by measuring the angle α of each cell from the horizontal
axis, or the axis parallel to the direction of radial flow. Because of the very high aspect
ratio of these cells in 3-D (typically greater than 10), the major axis of each cell was
readily identified and the angle calculated digitally from a manual trace. The cell angles,
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averaged in increments of 0.4 mm, were then plotted as a function of radial distance.
Since the average angle is 90° regardless of degree of alignment (as reflected in the
scatter plots in Fig. 4, fourth row panels), the variance of the average cell angle was used
as an indicator of the randomness of the cell orientation at a particular radial coordinate:
the larger the variance, the more random the cell orientation.
Computation of shear stress on cells under flow.
The shear stress on cells
subjected to interstitial fluid flow was estimated using a model by Wang and Tarbell for
estimating shear stresses resulting from transvascular flow on smooth muscle cells (29).
In this model, stress τ is estimated by τ= [Qµ/(2πrh)](K′)-1/2 (30). τ was calculated at the
inner and outer radii (ri and ro) to obtain estimates of the maximum and minimum
respectively, representing the range of shear stresses experienced by the cells undergoing
interstitial flow.
For flow on cells plated as 2-D monolayers, the radial profile of the wall shear
stress was derived from the velocity profile for steady radial flow between two parallel
disks with entrance effects ignored: τw=(3Qµ)/(πra2) where Q is the volumetric flow rate
and a is the distance between the top and bottom disks.
Results
Model features.
Our in vitro interstitial flow apparatus meets the following
criteria: (i) the culture system is supported against contraction and allows interstitial flow
without ECM compaction; (ii) the system is easily maintained over several days to allow
sufficient time to evaluate cell response; (iii) cell morphology and migration can be
monitored and visualized under microscopy in situ over a period of days, and (iv) the
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hydraulic resistance of the system can be continuously monitored to determine changes in
the most relevant mechanical property (i.e. flow resistance) and indirectly indicate matrix
integrity. In addition, the superficial flow velocity v and interstitial pressure P vary
across the radial distance of the gel, allowing a range of conditions to be examined and
compared in one experiment. All of these model features are demonstrated through the
following results.
Cells align perpendicular to interstitial flow.
Flow was induced through
fibroblast-populated collagen gels for 2-3 days at a volumetric flow rate of 10 µl/min.
Surprisingly, the fibroblasts displayed perpendicular alignment to the radial direction of
flow (Fig. 3A) and had a more noticeable spindle-shaped morphology than those in static
conditions. In contrast, the cells in static cultures retained a random orientation (Fig. 3B)
and appeared more branched in shape. In addition, we compared the effect of flow on
cells seeded in 3-D cultures with that on cells plated as a monolayer. The cells plated in a
monolayer were found to undergo regression under the same flow rate (Fig. 3C) or shear
stress (data not shown).
The radial design of our model allowed us to examine the dependence of cell
orientation on velocity due to the radial variations in velocity. Figure 4 compares the
profiles of cells undergoing interstitial flow with those in static conditions.
In the
cultures subjected to flow, velocity and pressure decreased radially (Fig. 4, first and
second row panels, respectively), while the hydrostatic pressure in the static gel (v=0
cm/s) was maintained at 1.082 cm H2O. On inspection of cell morphology (Fig 4., third
row panels), cells subjected to interstitial flow were perpendicularly aligned at higher
velocities but became more randomly oriented with increasing radial distance and thus
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decreasing bulk velocity. This was confirmed by the cell angle distributions (Fig. 4,
fourth row panels) and the increasing variance of the average cell angle with radial
distance (Fig. 4, fifth row panels). The estimated shear stress (30) on the cells in the
collagen matrix varied from 1.135 – 0.324 dynes/cm2 (at ri to ro respectively). Under
static conditions, cells were randomly oriented everywhere, as evidenced by the high and
consistent variance in angle across the radial distance.
Fibroblasts regress under 2-D fluid shear stress within 3 hours. The effects of
flow across a 2-D monolayer of fibroblasts were very different from those of interstitial
flow on 3-D cultured fibroblasts.
Within the same chamber and under the same
volumetric flow rate of 10 µl/min, cells regressed (Fig. 3C) within three hours. This flow
rate yielded a range of shear stresses on the surface of 0.048 – 0.014 dynes/cm2 from ri to
ro. We took time-lapsed images to determine the kinetics of regression and to ascertain
whether there was alignment of cells prior to regression (data not shown), but alignment
was not observed at any time prior to cell rounding. We also observed that with higher
flow rates, cells regressed even faster; thus, a comparison of cell morphology with similar
shear stress as that estimated for our interstitial flow studies – over an order of magnitude
larger - was not feasible. This was in contrast to static 2-D controls, which retained a
normal morphology throughout the experiment.
Darcy permeability reflects cell and matrix reorganization. K′ was measured by
monitoring the pressure drop across the chamber and comparing it with the imposed flow
rate according to Darcy’s law. When K' , the daily average of K′, was plotted against t,
we noticed that K' for the fibroblast-populated cultures showed an increase in the first
day (Fig. 5). However, K' eventually decreased before it leveled off at a steady value of
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about 0.25 x 10-6 cm2 by day 5.
On the other hand, the K ' for the acellular gel under
interstitial flow initially increased and leveled off at about 4.5 x 10-6 cm2.
Discussion
The effects of interstitial flow on cells are poorly understood despite the
importance of interstitial flow in tissue function. Here, we present a unique in vitro
interstitial flow model for cell-populated biological matrices along with novel
observations of cell alignment under interstitial flow within a 3-D matrix. Our interstitial
flow model maintains the following design criteria: (i) it can be visualized
microscopically in situ; (ii) it allows tracking of matrix remodeling via changes in
hydraulic conductivity; (iii) it does not contract macroscopically, and (iv) it can be
maintained over long periods of time since some cell and matrix organizational changes
are typically observed over a period of days to weeks. Although the data we present here
utilize dermal fibroblasts, which is a relevant system for investigating interstitial flow in
skin, the model can be easily extended to other cell systems such as vascular cells,
chondrocytes, or smooth muscle cells as well as with other matrices.
The main challenge to the creation of the model was to maintain the mechanical
integrity of the soft and highly compliant gel cultures undergoing interstitial flow without
compaction or fluid channeling around, rather than through, the gel. The problem was
complicated by cell contraction, which posed a serious challenge to our ability to monitor
the hydraulic conductivity of our cultures since gel contraction caused the sample to
detach, shrink, and reroute fluid flow. To address these problems, we used acid-treated
porous PE material to anchor the collagen gels on the lateral edges and surface
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modification of the glass surfaces on the top and bottom. Therefore, the collagen gel was
anchored in the radial direction by the PE rings and covalently bound to the top and
bottom glass surfaces to inhibit detachment while allowing visual observations in situ. In
addition, any gel contraction and fluid rerouting around the gel would significantly
decrease the hydraulic resistance and thus be reflected in the pressure manometer
readings. It should be noted that while compaction might be a relevant and desirable
feature for modeling interstitial flow through tissues such as cartilage (where compaction
drives the flow) or the arterial wall (where the pressure gradient that drives the flow also
expands the vessel lumen and compacts the wall), the boundaries of the interstitial space
between the capillary and lymphatics are fixed and thus the pressure gradients driving
this flow would tend to swell rather than compact the matrix.
We observed that interstitial cells (e.g. fibroblasts) respond to interstitial fluid
flow in a 3-D environment very differently than they do to pure shear flow when plated in
a 2-D monolayer, as in parallel plate flow chamber experiments. The fibroblasts under
interstitial flow align normal to the direction of flow whereas cells in static controls
retained a random orientation. In contrast, fibroblasts regressed into rounded structures
when they were plated as a 2-D monolayer and subjected to similar flow rates. Under
higher flow rates, such as to approach the estimated shear stress range in the 3-D cultures,
the cells regressed even more quickly, making a morphological comparison between 2-D
and 3-D cultures impractical. Furthermore, a direct comparison of shear stress between
the two systems may not be relevant since the mechanical environment is more complex
in the 3-D culture undergoing interstitial fluid flow due to the stresses on cell-ECM
attachments (e.g., shear stress on ECM fibers leads to ECM strain and thus cytoskeletal
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strain via integrins). Nonetheless, these results clearly demonstrate that 2-D flow studies
and 3-D static studies do not accurately reflect morphological behaviors of interstitial
cells that are mechanically stimulated in their natural 3-D configuration. It also suggests
that interstitial flow may affect how cells organize their environment.
The perpendicular alignment of the fibroblasts is consistent with observations in
other soft tissues where interstitial flow is significant, such as smooth muscle cells in the
arterial wall (12); however, it is important to note that circumferential stretch of the artery
wall is likely to be responsible for this alignment (18). One potential rationale for our
observations is that the cells realign in order to align the matrix to alter its mechanical
properties, as seen in matrix remodeling in other tissues in response to mechanical stress
(2, 12, 24). Interestingly, this perpendicular alignment undergoing interstitial flow is
different from other kinds of mechanical loading of fibroblast-populated collagen
matrices where parallel alignment of the cells to the direction of the applied load is
typically observed (6, 28).
The circumferential alignment of the fibroblasts appeared to occur in a critical
range of bulk velocities (Fig. 4).
The general trend we observed was that at high
superficial velocities, fibroblasts showed a rounded and regressed morphology; as bulk
velocity decreased, fibroblast morphology became more perpendicularly aligned and then
displayed random orientation at near-static conditions.
regressed at higher shear stress.
We hypothesized the cells
This was indirectly supported by morphological
observations of rounded fibroblasts at higher interstitial flow rates (e.g. 100 µl/min, data
not shown). In addition, the regressed cells were similar in morphology to those in the 2D fluid shear studies under much lower fluid shear rates. Furthermore, the anchoring of a
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fibroblast in a 3-D network of extracellular fibers provides more support against a
shearing force than if it were attached only on one side as in the 2-D monolayer; this may
account for the observations that fibroblasts could withstand higher shear stresses in 3-D
than in 2-D cultures.
One of the unique features about our model is that we can monitor changes in the
mechanical properties of the tissue culture under interstitial flow as an indirect online
monitor of matrix integrity and fluid channeling, since matrix composition and
architecture is the primary determinant of K′ (15). In our results, we noticed that K′ for
the cell-gel culture increased initially but later decreased and leveled off at a steady value
whereas K′ increased for the acellular gel but remained high. Although we have yet to
elucidate the mechanisms behind these observations, they may be indirect indications of
the fibroblasts’ ability to regulate their mechanical environment. The initial increase in
K′ may be due to fluid channeling within the gel, leading to a decrease in hydraulic
resistance. In cell-populated gels, the subsequent return of K′ to its lower initial value
may be the result of matrix synthesis and remodeling by the fibroblasts as well as perhaps
their realignment. Moreover, the measurements of K′ were consistently found to be in
the narrow range of 0.25 to 2.5 x 10-6 cm2, suggesting that while cells could affect the
hydraulic conductivity of the gel, they did not elicit drastic changes over the time period
of 5-7 days. This average K′ value is consistent with that obtained previously for type I
collagen (5).
In vivo measurements of K′ vary greatly due to location, method of measurement,
and state of hydration of the tissue, but are generally estimated to be in the range of 10-8
to 10-12 cm2 (15), with mouse tail skin recently estimated at 10-8 cm2 (27). Reconstituted
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collagen gels have a much higher value of K′ because collagen concentrations in cell
culture systems are by necessity very small (the maximum is about 3.5 mg/ml), and also
because reconstituted collagen gels do not have any specialized architecture nor are they
incorporated with other ECM molecules such as proteoglycans, which strongly decrease
K′ (15). Given this limitation, we could either mimic in vivo interstitial flow rates or
interstitial fluid pressure gradients in our system, but not both. Interstitial flow rates are
extremely difficult to measure in vivo, and estimates vary greatly due to measurement
technique and tissue type (31). On the other hand, interstitial fluid pressure – although
also difficult to measure and questionable – has been consistently found within a narrow
range, with average values in skin roughly within ± 2 cm H2O (32). Thus, our pressure
gradient driving the flow is physiologically relevant, even if the flow rates that result are
high due to the high K′. However, it is important to note that in vivo, K′ varies widely
according to hydration and thus level of swelling in a tissue. For example, Guyton
showed that changes in interstitial fluid pressure from –6.8 to +0.8 cm H2O led to
changes in K′ of over five orders of magnitude (9); this infers that interstitial fluid
velocity would also increase by five orders of magnitude for a given pressure gradient in
a slightly edematous tissue. Thus, because of the large variability of fluid balance
parameters found in vivo and particularly in edematous states, the range of interstitial
fluid pressure and average fluid velocity as well as the hydraulic resistance in our system
has clear physiological relevance.
In summary, the biological regulation of the interstitial fluid environment is
critical to understanding interstitial architecture in health and disease, including fluid
balance changes seen in edema. Our model can serve as a useful tool to study both short-
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and long-term effects of interstitial flow on cells and can easily be extended to examine
other cell systems and processes including those involved in angiogenesis, arterial wall
remodeling, and bone remodeling, to name a few. Other interests include the study of
fluid redistribution in the body due to gravitational effects such as deep sea diving and
weightlessness experienced in space. Using our model, we observed fibroblast alignment
perpendicular to interstitial flow.
Furthermore, the variability in the bulk Darcy
permeability K′ of the cultures suggested that interstitial flow was inducing matrix
reorganization towards some steady state, although the mechanisms responsible have yet
to be determined. These findings establish interstitial flow as having distinct effects on
cells different from those of 2-D shear flow, and our newly developed model is a useful
tool in understanding the biological regulation of interstitial fluid flow through soft
tissues.
Acknowledgements
The authors wish to thank Mr. Daniel Sedehi, Dr. Ranee Stile, Mr. Rick
Boardman, Dr. Matt Glucksberg and Dr. William Russin for invaluable technical
assistance and advice. This work was supported by the Whitaker Foundation and the
National Science Foundation.
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Figure Legends
Figure 1. Schematic of a microcirculatory fluid transport in a microvascular bed. Blood
flow in the capillaries (BC) brings fluid, which is then transported to cells in the
interstitium (I) via interstitial flow (arrows). Fluid is either then returned to the blood
microvessel or drained into the lymphatic microvessel (LC). Inset: Besides its role in
transport, interstitial flow also provides a mechanical environment directly on the
interstitial cells via the induction of shear stress (τ) and hydrostatic pressure (P), or
indirectly via cell-ECM signaling by imposing a strain (ε) and elastic stress (σ) on the
matrix fibers that the cells are attached to via integrin receptors.
Figure 2. Model and flow chamber set-up. A. The materials used for the chamber
consist of (a) silastic tubing, (b) functionalized glass slide, (c) porous PE rod, (e) porous
PE ring and (f) functionalized glass coverslip. Interstitial flow is delivered through the
cell populated collagen matrix (d); B. Photo of the assembled flow chamber; C. To
induce flow, the chamber (FC) is connected to a sterile media reservoir (R) via a
peristaltic pump (P) and a manometer (M).
Figure 3. Comparison of fibroblasts orientation in 3-D gels subjected to interstitial flow
(A) and in static controls (B) after 2 days with fluid-sheared fibroblasts plated as a subconfluent monolayer on a thin collagen layer after 3 hours (C). The arrows under
micrographs indicate the radial direction of the chamber. Taken at 100x magnification,
the images for the 3-D experiments (A and B) are inverted confocal micrographs of
maximum projections of flourescein phalloidin-stained cells in 150 µm thick sections
while the image for the monolayer shear stress experiment (C) is an inverted epifluorescence micrograph of Calcein AM-labeled cells. (Scale bar = 200 µm)
Figure 4. Comparison of typical radial sections from gels under interstitial flow and
static conditions.
One unique feature of the radial design is that the variation of
superficial flow velocity (v=Q/(2πrh)) and pressure (P=Po + [Qµ ln(ro/ri)]/[2πhK′(t)])
along the radial distance (r*=(r-ri)/(ro-ri)). Velocity (1st row panels) and pressure profiles
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(2nd row panels) with cell orientation α (4th row panels) and variance data Var( α ) (5th
row panels) were plotted against the radial distance (arrows). The inverted confocal
images (3rd row panels) are maximum projections of 150 µm thick sections of gels with
flourescein phalloidin-stained cells taken at 100x magnification.
Figure 5. One of the unique features of the flow model is that we can monitor Darcy
permeability changes over time. Variability in the average bulk Darcy permeability data
for the fibroblasts (♦) and acellular gels (▲), averaged daily and presented in the graph
as K' , suggests that interstitial flow was inducing matrix reorganization towards some
optimal state.
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Figure 1
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Figure 2
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Figure 3
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Figure 4
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Figure 5